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*
Department of Pediatrics, University Hospital, Nijmegen, The
Netherlands.
Department of Pathology, University Hospital, Nijmegen, The
Netherlands.
§
Department of Hematology, University Hospital, Nijmegen, The
Netherlands.
Gaubius Laboratory TNO Prevention and Health, Leiden, The
Netherlands.
Correspondence to Dr. Victor W. M. van Hinsbergh, Gaubius Laboratory TNO-PG, Zernickedreef 9, P.O. Box 2215, 2301 CE Leiden, The Netherlands. Fax: + 31-71-5181904; E-mail: vwm.vanhinsbergh{at}pg.tno.nl
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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(TNF-). FMVEC displayed strong
binding of VT and high susceptibility to VT under basal conditions, which made
them suitable for the study of VT-induced apoptosis without TNF-
interference. On the basis of functional (flow cytometry and
immunofluorescence microscopy using FITC-conjugated annexin V and propidium
iodide), morphologic (transmission electron microscopy), and molecular
(agarose gel electrophoresis of cellular DNA fragments) criteria, it was
documented that VT induced programmed cell death in microvascular endothelial
cells in a dose- and time-dependent manner. Furthermore, whereas partial
inhibition of protein synthesis by VT was associated with a considerable
number of apoptotic cells, comparable inhibition of protein synthesis by
cycloheximide was not. This suggests that additional pathways, independent of
protein synthesis inhibition, may be involved in VT-mediated apoptosis in
microvascular endothelial cells. Specific inhibition of caspases by
Ac-Asp-Glu-Val-Asp-CHO, but not by Ac-Tyr-Val-Ala-Asp-CHO, was accompanied by
inhibition of VT-induced apoptosis in FMVEC and TNF--treated GMVEC.
These data indicate that VT can induce apoptosis in human microvascular
endothelial cells. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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1—4Gal in a terminal
position (5). The biologic
activity of the toxins is thought to involve inhibition of overall protein
synthesis through enzymatic inactivation of the 60S ribosomal units
(6). Induction of apoptosis has
also been reported for a variety of cell types
(7,8,9),
including renal tubule-derived epithelial cells
(10). The presence of
apoptotic endothelial cells in the glomeruli of kidney biopsy specimens from
three patients with the epidemic form of HUS was recently reported
(11). However, whether VT can
directly induce apoptosis in endothelial cells has not been studied. Apoptosis
(programmed cell death) is a highly regulated process characterized by
morphologic, functional, and molecular features
(12,
13). Apoptosis occurs not only
under physiologic conditions, such as embryogenesis, morphogenesis, and other
processes of tissue formation and cell renewal, but also under pathologic
conditions. A number of physical and chemical agents, including a variety of
toxins, have been reported to induce apoptosis
(7,8,9,10,
14). Apoptosis can be induced
by several activation pathways, which cause the activation of procaspases into
active caspases (13,
15). These activities converge
in the activation of the so-called death caspases (caspases 3, 6, and 7),
which initiate an irreversible process that leads to DNA fragmentation, cell
detachment, and death (15,
16). The presence of caspase
1-like activity and caspase 3 has been demonstrated in human umbilical vein
endothelial cells (HUVEC)
(17).
In vitro studies using HUVEC and glomerular microvascular
endothelial cells (GMVEC) have indicated that VT susceptibility requires
additional stimulation by inflammatory mediators for induction of a
sufficiently large number of specific VT receptors on these cells
(5,
18). Because these mediators
may themselves induce both stimulators and inhibitors of apoptosis in
endothelial cells
(19,20,21),
it would be extremely difficult to discriminate between direct and indirect
effects of VT on apoptosis in tumor necrosis factor-
(TNF-)-stimulated GMVEC. In our studies, we observed that unstimulated
human foreskin microvascular endothelial cells (FMVEC) are very sensitive to
VT cytotoxicity in vitro. These cells provided a means to evaluate
whether VT could induce apoptosis in human microvascular endothelial cells
without prior stimulation with TNF-. Using morphologic, functional, and
molecular criteria, we demonstrate that VT induces apoptosis in these cells
and also induces apoptosis in TNF--stimulated GMVEC. Using specific
inhibitors, we investigated whether caspase 3 (CPP32, apopain) and caspase
1-like [interleukin-1ß (IL-1ß)-converting enzyme (ICE)-like]
proteases play a role in VT-induced apoptosis.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Endothelial Cell Cultures
FMVEC were isolated and purified according to methods previously described
by Davison et al.
(23) and Voyta et al.
(24). FMVEC were seeded on
gelatin (1%; Fluka Biochemicals, Buchs, Switzerland)-coated, six-well plates
(Costar, Cambridge, MA) and cultured in M199 (BioWhittaker, Walkersville, MD)
supplemented with 10% (vol/vol) human serum (local blood bank), 10% (vol/vol)
newborn calf serum (NBCS) (Life Technologies, Grand Island, NY), 2 mM
L-glutamine (ICN Biomedicals, Zoetermeer, The Netherlands), 5 U/ml heparin
(Leo Pharmaceutical, Weesp, The Netherlands), 100 IU/ml penicillin/0.1 mg/ml
streptomycin (Yamanouchi Pharma B.V., Leiderdorp, The Netherlands), and 150
mg/liter crude endothelial cell growth factor [extracted from bovine brains as
described by Maciag et al.
(25)]. Cells were subcultured
with mild trypsinization (3 to 5 min), using
trypsin/ethylenediaminetetraacetate (EDTA) (0.5 g/liter trypsin, 1:250, and
0.2 g/liter EDTA; Life Technologies), after which the cells were replated with
a split ratio of 1:3. For experiments, FMVEC were used after nine to 11
passages.
GMVEC were isolated and purified as previously described by van Setten et al. (18). HUVEC were isolated from umbilical cords according to the method described by Jaffe et al. (26). GMVEC and HUVEC were used after eight to 10 and two to four passages, respectively.
All endothelial cells used displayed the presence of von Willebrand factor,
platelet endothelial cell adhesion molecule-1, and V- and E-cadherin at all
passages used. No immunoreactivity to the anticytokeratin 20 antibody or the
anti--smooth muscle actin antibody was observed, excluding the
possibility of contaminating epithelial and mesangial cells, respectively
(18).
Cytotoxicity Assays
Microvascular endothelial cells were cultured in complete medium on
gelatin-coated, 24-well plates and were grown until confluence. Five days
after reaching confluence, cells were preincubated with or without TNF-
(10 ng/ml) for 24 h. The next day, the medium was aspirated and fresh medium
with different concentrations of VT was added. VT was diluted in medium with
20% fetal calf serum instead of 10% NBCS and 10% human serum, because previous
studies indicated that NBCS and human serum may have neutralizing activity
against VT. After 24 h, the cells were washed with phosphate-buffered saline
(PBS) and released with trypsin/EDTA. Viable, trypan blue-excluding cells were
then counted with a hemocytometer.
Protein Synthesis
Protein synthesis was assessed by assaying the incorporation of
35S-labeled methionine (0.25 µCi/ml complete medium) into
35S-proteins during 24-h incubations with different concentrations
of VT. After incubation, the cells were washed and 35S-labeled
cellular proteins and 35S-labeled proteins present in the medium
were precipitated with the addition of TCA (10 and 20%, respectively).
Precipitated proteins were dissolved in 0.3 ml of 0.3 M NaOH, 60 µl of 1.5
M HCl was added, and precipitated radioactivity was counted in a liquid
scintillation counter
(18,27).
Glycolipid Extraction and Thin Layer Chromatography
FMVEC were cultured in complete medium on gelatin-coated, six-well plates
(Costar) and were grown to confluence. Highly confluent cells were exposed to
complete medium alone or complete medium supplemented with TNF- (10
ng/ml). After 24 h, the cells were trypsinized and washed three times with
ice-cold PBS. Subsequently, glycolipids were extracted, separated, and assayed
for 125I-VT binding, as described previously
(18,27).
Thin layer chromatograms were analyzed with a Fuji BAS 1000 PhosphorImager
(Fuji Photo Film Co., Tokyo, Japan).
Apoptosis Assays
Fluorescence-Activated Cell Sorting Analyses. FMVEC, HUVEC, and
GMVEC were grown on gelatin-coated, 12-well plates (Costar) until they reached
confluence. Highly confluent HUVEC and GMVEC were prestimulated with
TNF- (10 ng/ml) for 24 h, to upregulate VT receptors and induce VT
susceptibility
(18,27).
Because FMVEC display VT susceptibility under basal conditions, these cells
were not pretreated with TNF-. Subsequently, unstimulated FMVEC and
TNF--stimulated HUVEC and GMVEC were incubated for 4 or 24 h with
complete medium, with complete medium supplemented with various concentrations
of VT (0.1 pM to 10 nM), B-subunit (5 to 130 nM), or cycloheximide (CHX) (0.01
to 1 µg/ml), or with M199 alone. After the incubation period, detached
cells were collected and pooled with trypsinized adherent cells. Cells were
centrifuged at 200 x g for 5 min at 4°C, and the
supernatant was removed. The cells were then washed with ice-cold binding
buffer (10 mM Hepes, [pH 7.4], 150 mM NaCl, 5 mM KCl, 1 mM MgCl2,
1.8 mM CaCl2), as previously described by Koopman et al.
(28), supplemented with 0.1%
bovine serum albumin (BSA) (ICN Biomedicals, Zoetermeer, The Netherlands).
Cells were resuspended in 500 µl of binding buffer with 0.1% BSA, of which
445 µl was transferred into a tube suitable for fluorescence-activated cell
sorting (FACS) analysis. Five microliters of FITC-conjugated annexin V (2
µg/ml, diluted in binding buffer with 0.1% BSA; Bender Medsystems, Vienna,
Austria) and 50 µl of propidium iodide [100 µg/ml, diluted in RPMI DM
(Flow Inc.) supplemented with 5% fetal calf serum and 2 mM CaCl2;
Sigma] were added, after which the cell suspension was gently mixed and
incubated in the dark on ice for 10 min
(28,29).
Samples were assayed for viable, apoptotic, and necrotic cells by FACS
analysis (Coulter 7 Epics 7 XL-MCL, Beckman Coulter Inc., Mijdrecht, The
Netherlands). Necrotic cells were defined as cells demonstrating positive
staining for both FITC-conjugated annexin V and propidium iodide. Viable cells
were not positive for either FITC-conjugated annexin V or propidium iodide.
Apoptotic cells were defined as cells exhibiting positive staining for
FITC-conjugated annexin V and negative staining for propidium iodide.
Fluorescence was measured on a double-parameter histogram, using logarithmic
scales. For each tube, 5000 events were analyzed.
To assess the inhibition of the apoptosis-inducing effect by reversible inhibitors, FMVEC, cultured under identical conditions, were preincubated for 1 or 24 h with various concentrations of ICE inhibitor I [Ac-Tyr-Val-Ala-Asp-CHO (YVAD-CHO); Calbiochem, San Diego, CA] or CPP32/apopain inhibitor [Ac-Asp-Glu-Val-Asp-CHO (DEVD-CHO); Calbiochem] (30,31). After preincubation with these inhibitors, cells were rinsed once with M199, after which the cells were exposed for 16 h to complete medium or complete medium supplemented with VT (1 nM), in the presence of the same reversible inhibitor. Apoptosis was evaluated by FACS analysis as described previously. To detect the presence of caspase 3, caspase 6, or caspase 7 in FMVEC and GMVEC, Western blot analysis was performed according to established procedures (31), using primary antibodies against human CPP32/p20 (caspase 3) (N-19), Mhc2/p20 (caspase 6) (K-20), or ICE-LAP3 (caspase 7) (C-18) (Santa Cruz Biotechnology, Santa Cruz, CA) and a horseradish peroxidase-conjugated anti-goat IgG antibody (diluted 1:5000 in PBS with 0.5% BSA and 0.05% Tween-20; Nordic Immunology, Tilburg, The Netherlands) as a secondary antibody.
Fluorescence Microscopy Using Triple Staining of Cell Monolayers. Cells were seeded onto gelatin-coated glass coverslips, as described previously (18), and were grown to confluence. Highly confluent FMVEC were exposed to complete medium alone or complete medium supplemented with various concentrations of VT. After an incubation period of 16 h, adherent cells were stained with 500 µl of Hoechst 33342 (1 µg/ml in complete medium; Sigma) for 15 min at 37°C. Subsequently, cells were rinsed once with M 199 and incubated, on ice, with 500 µl of complete medium containing FITC-conjugated annexin V (1 µg/ml) and propidium iodide (1 µg/ml). Cells were analyzed by fluorescence microscopy within 15 min, using filters of 365, 490, and 560 nm for Hoechst 33342, FITC-conjugated annexin V, and propidium iodide staining, respectively. Definitions of viable, apoptotic, and necrotic cells were similar to those described for the FACS analyses.
DNA Fragmentation Analyses. FMVEC were grown in complete medium on gelatin-coated, six-well plates until they reached confluence. Highly confluent cells were exposed to complete medium or complete medium supplemented with various concentrations of VT or B-subunit. After 16 h of incubation, detached cells were collected on ice and pooled with trypsinized adherent cells. Cells were subjected to centrifugation at 200 x g for 5 min at 4°C, and the pellets were washed twice with ice-cold PBS. Subsequently, cells were lysed with 10 mM Tris (Boehringer Mannheim, Mannheim, Germany), pH 8.0, 10 mM EDTA (Life Technologies), 0.5% Triton X-100 (Boehringer Mannheim), as previously described by Bissonnette et al. (32). To separate fragmented DNA from intact chromatin, cells were centrifuged at 15,000 x g for 20 min at 4°C. Soluble fragmented DNA was transferred into a new tube and treated with RNase A (100 µg/ml; Sigma) for 1 h at 37°C, followed by treatment with proteinase K (200 µg/ml; Sigma) in 1% sodium dodecyl sulfate (Merck, Darmstadt, Germany) for 2 h at 50°C. DNA was precipitated with 0.1 volume of 3 M sodium acetate and 2 volumes of 96% ethanol for 16 h, after which the samples were centrifuged at 15,000 x g for 10 min at 4°C. Subsequently, DNA pellets were air-dried and dissolved in Tris-EDTA solution, pH 8.0 (10 mM Tris-HCl and 0.1 mM EDTA, both at pH 8.0). Loading buffer was added to each sample, and electrophoresis was performed at 80 V for 1.5 to 2 h on a 1.5% agarose gel containing 0.4 µg/ml ethidium bromide (Merck), with 1 x Tris/borate/EDTA running buffer (22.5 mM Tris-borate, 0.5 mM EDTA, pH 8.0). DNA was observed under ultraviolet light (350 nm) and photographed. DNA size calibration was performed using a 100-bp marker (Pharmacia, Roosendaal, The Netherlands).
Transmission Electron Microscopy. To evaluate the apoptosis-inducing effect of VT on adherent endothelial cells by transmission electron microscopy, FMVEC were cultured in complete medium on gelatin-coated, 12-well plates. After they reached confluence, the cells were exposed to control medium or control medium supplemented with various concentrations (0.1 pM to 10 nM) of VT. After 16 h of incubation, detached cells were pooled with trypsinized adherent cells. Cells were pelleted and fixed for 2 h at room temperature with 2.5% glutaraldehyde in cacodylate buffer. The cells were then carefully rinsed with 0.1 M cacodylate buffer for 30 min, after which the cells were immersed and pelleted in 15% gelatin. After immersion, alternate fixation with 1% OsO4 in cacodylate buffer was performed for 30 min. Cells were then selectively dehydrated in graded ethanol and embedded in Epon 812. Thin sections were cut with an ultramicrotome and stained with uranyl acetate and lead citrate. Sections were examined with a Jeol 1200 EX electron microscope (Jeol Europe bv., Schilphol-Rijk, The Netherlands).
Additional Analytic Procedures
To determine whether the cultured cells exhibited signs of endogenous
activation, the expression of monocyte chemoattractant protein-1 (MCP-1)
(33), urokinase-type
plasminogen activator (u-PA)
(34), and vascular cell
adhesion molecule-1 (VCAM-1)
(18) in culture supernatants
(u-PA and MCP-1) or on the cell surface (VCAM-1) was assessed according to
established procedures.
Statistical Analyses
The statistical significance of differences between groups was analyzed by
ANOVA followed by Bonferroni's modified t test. Differences were
considered significant at P < 0.05 (two-sided).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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or IL-1 to
become sensitive to VT. Exposure of unstimulated FMVEC to various
concentrations of VT (0.01 fM to 1 nM) for 24 h produced
concentration-dependent cell toxicity and inhibition of overall protein
synthesis (Figure 1). This high
sensitivity to VT was related to high levels of specific VT binding to these
cells, as assayed in binding experiments with 125I-VT. Thin layer
chromatographic analyses of cellular neutral glycolipids extracted from
unstimulated FMVEC revealed strong binding of 125I-VT to glycolipid
species in the Gb3 region (Figure
2), with an additional increase after stimulation with
TNF-.
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To investigate whether FMVEC had spontaneously acquired the activation
phenotype that is induced by TNF- or IL-1, we measured the levels of
several compounds that are induced in endothelial cells by TNF- and
IL-1. Values for VCAM-1 (not detectable), MCP-1 (<0.3 ng/ml), and u-PA (0.2
ng/ml) in unstimulated FMVEC were essentially identical to those found in
unstimulated GMVEC and HUVEC, and values increased during 24-h exposure to
TNF- (VCAM-1 clearly expressed; MCP-1, >2000 ng/ml; u-PA, 1.7 ng/ml;
average of three independent determinations).
VT Induction of Apoptosis in FMVEC
To evaluate whether apoptosis is involved in VT-mediated FMVEC cell death,
we initially performed FACS analyses using a dual-staining method with
FITC-conjugated annexin V and propidium iodide. This method discriminates
among viable, apoptotic, and necrotic cells
(28,29)
(see the Materials and Methods section for details).
Figure 3 presents
representative cytograms for control
(Figure 3A) and VT-treated
(Figure 3B) FMVEC. Whereas in
control cells the numbers of apoptotic and necrotic cells were rather small (3
and 5%, respectively), VT-treated (0.1 nM for 24 h) cells demonstrated an
impressive increase in apoptotic cells (40.7 ± 4.5%, n = 3)
and a slight increase in necrotic cells (12.3 ± 2.9%, n =
3).
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The apoptosis-inducing effect of VT in FMVEC was time-and concentration-dependent (Figure 4). The apoptosis-inducing effect of VT was observed as early as 4 to 6 h after the beginning of the VT incubation, with an increase in the number of apoptotic cells after 24 h of VT exposure. The threshold dose for VT to induce apoptosis was 0.1 pM. Identical results were obtained when VT-1, VT-2, and VT-2c were compared (Table 1). The B-subunit of VT induced apoptosis only at very high concentrations. Exposure of the cells to 50 and 130 nM B-subunit for 24 h resulted in 29 ± 8% and 40 ± 8% apoptotic cells (mean ± SD of two independent experiments with two different donors), respectively. The percentages of necrotic cells were 9 ± 0.2% and 11 ± 2%, respectively.
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VT-induced apoptosis in FMVEC was confirmed by direct observation, using a triple-staining method. As shown in Figure 5, highly confluent control and VT-exposed cells were stained with Hoechst 33342 (Figure 5, A and B) or FITC-conjugated annexin V and propidium iodide (Figure 5, C and D). Staining of control cells with Hoechst 33342 resulted in homogeneous diffuse nuclear staining (Figure 5A). After incubation of FMVEC with VT (10 nM) for 16 h, some of the adherent cells displayed condensed and fragmented nuclear chromatin, a characteristic feature of apoptosis (Figure 5B, arrows). Whereas almost all (>99%) control cells were negatively stained with annexin V and propidium iodide (Figure 5C), indicating that they were viable, a major fraction of VT-treated FMVEC demonstrated positive staining with annexin V and negative staining with propidium iodide, indicating apoptosis (Figure 5D, arrows). A minority of VT-exposed cells exhibited intracellular staining with annexin V and propidium iodide, which is characteristic of necrotic cells (Figure 5D, star).
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DNA Fragmentation Analysis and Ultrastructural Morphologic Features
of VT-Exposed FMVEC
Observations made using FACS analysis and fluorescence microscopy were
extended by DNA fragmentation analyses. Endonuclease-induced cleavage of
nuclear DNA into mono-and oligonucleosomal fragments with approximate
molecular sizes of multiples of 180 nucleotides is a characteristic feature of
programmed cell death (35).
Figure 6 demonstrates the
electrophoretic patterns of DNA extracted from control and VT
holotoxin-treated FMVEC. Whereas DNA from control cells remained intact
throughout the 16-h incubation period
(Figure 6, lane C), DNA from VT
holotoxin (0.1 fM to 1 nM)-exposed cells clearly exhibited the DNA
"ladder" pattern of multiples of 180-bp fragments. Exposure of the
cells to 130 nM VT B-subunit induced a similar DNA ladder pattern (data not
shown).
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In addition, the apoptosis-inducing effect of VT on FMVEC was studied by transmission electron microscopy. At the ultrastructural level, apoptotic cells exhibit distinctive alterations, primarily of the nucleus, which can be discriminated from events of necrosis (35). Representative micrographs of control and VT-treated FMVEC are presented in Figure 7. The morphologic features of control FMVEC in culture were maintained; cells exhibited nuclei with normal heterogeneous chromatin and a nuclear envelope, normal intact organelles, and normal plasma membranes (Figure 7A). In contrast, FMVEC exposed to VT (1 nM) for 16 h exhibited varying degrees of nuclear condensation (Figure 7, B and C), and even cellular fragmentation was noted (Figure 7D). Furthermore, we observed abundant cytoplasmic vacuolization and the presence of apoptotic bodies; however, cellular organelles of VT-treated FMVEC appeared normal. Although the integrity of the plasma membrane remained intact, blebbing of the plasma membrane occurred in VT-exposed FMVEC. These morphologic changes are indicative of apoptosis and are clearly different from those exhibited by cells undergoing necrosis.
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Comparison of Apoptosis Induced by VT and CHX
To evaluate whether the induction of apoptosis by VT merely reflects
inhibition of protein synthesis, we compared the effect of VT with that of the
protein synthesis inhibitor CHX. For proper comparisons, we determined which
CHX concentration inhibited protein synthesis to a similar extent, compared
with VT (0.1 pM), in wells cultured in parallel. Because 0.1 µg/ml CHX and
0.1 pM VT both inhibited protein synthesis by 30%, their effects on apoptosis
induction were compared. Only VT induced apoptosis (14 ± 3% for VT
versus 2 ± 1% for CHX; mean of two independent experiments).
These data indicate that VT is a more potent inducer of apoptosis than is CHX
at comparable potencies for protein synthesis inhibition.
Apoptosis of TNF--Pretreated GMVEC and HUVEC after Exposure to
VT
The effect of VT on the induction of apoptosis among GMVEC and HUVEC was
subsequently investigated by FACS analysis, using a dual-staining method with
FITC-conjugated annexin V and propidium iodide. VT did not induce significant
cell death among untreated GMVEC or HUVEC (data not shown). However, when
GMVEC or HUVEC were preincubated with TNF- for 24 h, 4- to 6-h exposure
of the cells to VT (0.1 to 10 nM) induced a dose-dependent increase in the
number of apoptotic cells (Figure
8). Although no further increase in the percentage of apoptotic
cells was observed after 24 h of VT exposure, the percentage of necrotic cells
increased significantly.
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Effects of Caspase Inhibitors on VT-Induced Apoptosis of FMVEC and
GMVEC
Caspases such as caspase 1 (ICE) and caspase 3 (CPP32) have been implicated
in the complex cascade of events that results in apoptosis
(15). To determine whether
these cysteine proteases play a role in VT-mediated apoptosis, DEVD-CHO and
YVAD-CHO, specific inhibitors that discriminate between different caspases
(inhibiting caspase 3 and caspase 1-like proteases, respectively), were used.
FMVEC incubated with DEVD-CHO (3 to 300 µM) demonstrated dose-dependent
inhibition of VT-induced apoptosis. The threshold dose to decrease the
percentage of apoptotic cells was 30 µM; a maximal response of 60 to 70%
inhibition of VT-mediated apoptosis was obtained with 300 µM DEVD-CHO
(Tables 2 and
3). Similarly, DEVD-CHO partly
inhibited VT-induced apoptosis in TNF--exposed GMVEC
(Table 2). Administration of
YVAD-CHO (3 to 300 µM) did not result in significant inhibition of
VT-induced apoptosis in FMVEC (Table
2). Western blotting of cell extracts demonstrated that both FMVEC
and unstimulated and TNF--stimulated GMVEC contained caspase 3, whereas
caspases 6 (Mhc-2) and 7 (ICE-LAP, Mhc-3) could not be detected (data not
shown). These data are consistent with a possible involvement of caspase 3 in
VT-induced apoptosis in human microvascular endothelial cells.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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and in unstimulated FMVEC and probably involves the activation of caspase
3. Endothelial cell damage of predominantly glomerular capillaries is a characteristic feature in the pathogenesis of the epidemic form of HUS (2). From a pathogenetic point of view, it is generally assumed that VT is potentially cytopathic for endothelial cells. Several in vitro observations have indicated that priming of the endothelial cells by inflammatory mediators is required for VT cytotoxicity. These mediators cause an increase in VT susceptibility via enhancement of the number of VT-binding receptors (18,27). The mechanisms reported for VT-induced cell death in a variety of cell types, including endothelial cells, involve A-subunit-dependent inhibition of overall protein synthesis through the enzymatic inactivation of 60S ribosomal units (6). Observations in Vero cells (7,9), Burkitt's lymphoma cells (8), and renal tubule-derived epithelial cells (10) have indicated that VT also induces programmed cell death. VT-mediated apoptosis in cultured endothelial cells has not yet been reported.
Here we demonstrate that VT induces apoptosis in TNF--treated GMVEC
and HUVEC. Because these cells require prestimulation with TNF- to
acquire increased numbers of VT receptors and high sensitivity to VT
(18,27),
a direct effect of VT on apoptosis is difficult to establish. On one hand,
TNF- itself has been reported to induce apoptosis in lymphocytes
(36) and bovine endothelial
cells
(19,20).
On the other hand, TNF- has been reported to induce the expression of
A1, a Bcl-2 homolog that is known to block programmed cell death in human
endothelial cells (21).
Subsequent exposure to protein synthesis inhibitors, including VT, may cause a
decrease in the expression of this apoptosis-blocking homolog, resulting in
the induction of apoptosis. It would thus be extremely difficult to establish
whether VT directly or indirectly induces apoptosis. In our studies, we
observed that unstimulated FMVEC demonstrated high levels of VT binding and
high VT sensitivity, with only minor increases after stimulation with the
inflammatory mediator TNF-. The reason why unstimulated FMVEC express
many VT receptors (Gb3 molecules) is unknown. It is not a reflection of
general TNF--like activation of the cells but may be attributable to
reduced catabolism of Gb3 or markedly reduced conversion of Gb3 to Gb4. It is
also not known whether the large number of VT receptors in these cells
represents an in vitro feature of these cells or also occurs in
vivo. Nevertheless, in various assays with these cells, we obtained
evidence that VT can directly induce apoptosis in endothelial cells, even
without prior exposure to TNF-. These observations make it likely that
VT can also directly induce apoptosis in TNF--treated GMVEC.
Previous observations indicated that certain cell populations undergo apoptosis when exposed to inhibitors of protein synthesis (37,38,39). These data may indicate that VT-mediated apoptosis in FMVEC results from the inhibitory effect of VT on protein synthesis. Interestingly, whereas partial inhibition of protein synthesis by VT was associated with a considerable number of apoptotic cells, comparable inhibition of protein synthesis by CHX, which is known to block the peptidyl transferase reaction on ribosomes, was not. The finding that high concentrations of the B-subunit of VT alone induced similar features of apoptosis in FMVEC, compared with the holotoxin, further suggests that additional pathways, independent of protein synthesis inhibition, may contribute to VT-mediated apoptosis. The latter finding is closely related to observations made by Mangeney et al. (8), who reported that the B-subunit of VT induced apoptosis in Burkitt's lymphoma cells. However, in Vero cells, which, like FMVEC, are highly susceptible to VT, only VT holotoxin and not the B-subunit induced programmed cell death (40). This leaves open the possibility that the effect of high concentrations of the B-subunit occurs in addition to the effect of VT itself.
In our studies, we used primarily VT-2c, which is structurally and functionally related to VT-1 and VT-2 (22). VT-1, VT-2, and VT-2c are associated with the epidemic form of HUS in childhood, but the former two are more commonly associated with human disease than is VT-2c. As pointed out in our comparative studies, the apoptosis-inducing effect of VT-2c is similar to that of VT-1 and VT-2. Identical effects of different forms of VT were previously demonstrated for inhibition of protein synthesis (18) and cell proliferation (33).
Cell death by apoptosis involves the activation of caspases, followed by
caspase-mediated proteolysis and nucleosomal DNA fragmentation. Similar to the
coagulation system, a cascade of proteolytic activations occurs, beginning
with the activation of procaspases with a long prodomain and converging in the
activation of procaspases with a short prodomain
(15,41).
These latter caspases, i.e., caspases 3, 6, and 7, are also referred
to as death caspases, because they initiate processes that irreversibly cause
cell death. Our observation that a peptide aldehyde inhibitor of caspase 3 and
related caspases, such as caspases 7 and 8, and not an inhibitor of caspase 1
inhibited VT-mediated apoptosis in FMVEC and TNF--stimulated GMVEC
indicates that VT-mediated apoptosis is linked to activation of the former
caspases and not that of caspase 1. Caspase 3 was present in relatively large
quantities in both FMVEC and GMVEC, in agreement with previous observations in
HUVEC
(17,31,42).
Whereas caspase 3 was demonstrated in cultured endothelial cells, it could not
be demonstrated in quiescent endothelial cells in vivo
(43). However, the induction
of caspase 3 in pathologic conditions in vivo has not yet been
evaluated. Future studies may provide further insight into the complex
molecular mechanisms underlying the apoptosis-inducing effect of VT in
endothelial cells. In particular, it may be of interest to study the roles of
glycolipid breakdown products, such as ceramide and sphingosine, and their
phosphorylation products. These products have been reported to be involved in
programmed cell death in other cell types
(44).
Apoptosis is thought to represent an important defense mechanism in the event of cell damage. In contrast to necrosis, this type of cell death is usually followed by the removal of unwanted cells without the induction of an inflammatory response. Therefore, apoptosis of endothelial cells may allow uncompromised restoration of the endothelial cell layer. However, inflammation seems to precede VT-induced cell death in GMVEC. Therefore, we hypothesize that apoptosis is part of a complex cascade that takes place inside the glomerular capillaries and finally leads to kidney failure. Because apoptotic cells have been reported to be procoagulant (45), they may contribute to thrombotic events in HUS and thus are likely to amplify glomerular endothelial cell injury.
Mitra et al. (46) reported that plasma from four patients with thrombotic thrombocytopenic purpura (which is related to HUS) could induce apoptosis in restricted lineages of microvascular endothelial cells. This could not be demonstrated with plasma from one patient with HUS associated with diarrhea. However, whether VT is present at sufficient levels in the plasma of patients with HUS remains uncertain. Recently, Karpman et al. (11) reported for the first time the presence of apoptotic endothelial cells in the glomeruli of kidney biopsy specimens from three patients with the epidemic form of HUS. The occurrence of apoptotic features in the epidemic form of HUS may be underestimated. Descriptions of the pathologic features of HUS included primarily autopsy or biopsy findings for kidney specimens obtained at an advanced stage of the disease. In such samples, necrosis secondary to apoptosis, as well as phagocytosis of small numbers of apoptotic cells by macrophages and adjacent cells, might have occurred. In addition, most pathologic studies used light microscopy, with which it is difficult to detect and distinguish apoptotic and necrotic cells. Therefore, advanced techniques applied early in the course of the disease are needed to specifically detect apoptotic cells in tissue samples.
We conclude from this study that VT can induce apoptosis in human microvascular endothelial cells by a mechanism that possibly involves caspase 3. These observations may provide further insight into the pathogenesis and pathophysiologic features of VT-mediated microvascular endothelial cell damage in HUS.
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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on verocytotoxin cytotoxicity in purified human
glomerular microvascular endothelial cells. Kidney Int51
: 1245-1256,1997[Medline]
: Modulation by
inhibitors of protein synthesis. Exp Cell Res214
: 584-594,1994[Medline]
. J Immunol 149:2089
-2094, 1992[Abstract]
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), control GMVEC (
), and
tumor necrosis factor-
)
were incubated with different concentrations of VT (0.01 fM to 10 nM). After
24 h, the number of viable cells (A) and the incorporation of
[35S]methionine into proteins (B) were determined as described in
Materials and Methods. Data are expressed as the mean ± SD of two
independent experiments, with two different donors [percentage of control (%
of C)].
) incubation period. Detached and adherent cells were obtained
and assayed for the occurrence of apoptosis by FACS analysis, as described in
Materials and Methods. Data are expressed as the mean ± SD of three
independent experiments with cells from two different donors.
, 1 nM; 
, 10 nM) for 5 h.
Subsequently, detached cells and trypsinized adherent cells were pooled and
assayed for the presence of apoptotic cells by FACS analysis, as described in
Materials and Methods. (A and C) Data are expressed as the mean ± SD of
four (HUVEC) (A) or three (GMVEC) (C) independent experiments with cell
cultures from different donors. (B and D) Representative flow cytometric
analyses of control and VT (10 nM)-treated, TNF-